BioMed Research International

Volume 2014 (2014), Article ID 135218, 26 pages

http://dx.doi.org/10.1155/2014/135218

Research Article

Phylogenetic Analysis of Entomoparasitic Nematodes, Potential Control Agents of Flea Populations in Natural Foci of Plague

E. I. Koshel,¹ V. V. Aleshin,2,3,4 G. A. Eroshenko,¹ and V. V. Kutyrev¹

¹Russian Research Anti-Plague Institute “Microbe”, Saratov 410005, Russia

²Belozersky Institute of Physical-Chemical Biology, Lomonosov Moscow State University, Moscow 119991, Russia

³Institute for Information Transmission Problems, Russian Academy of Sciences, Moscow 127994, Russia

⁴National Research Institute of Physiology, Biochemistry, and Nutrition of Farm Animals, Russian Academy of Agricultural Sciences, Kaluga Region, Borovsk 249013, Russia

Received 28 February 2013; Revised 5 June 2013; Accepted 22 October 2013; Published 3 April 2014

Academic Editor: Vassily Lyubetsky

Copyright © 2014 E. I. Koshel et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Abstract.

Entomoparasitic nematodes are natural control agents for many insect pests, including fleas that transmit Yersinia pestis, a causative agent of plague, in the natural foci of this extremely dangerous zoonosis. We examined the flea samples from the Volga-Ural natural focus of plague for their infestation with nematodes. Among the six flea species feeding on different rodent hosts (Citellus pygmaeus, Microtus socialis, and Allactaga major), the rate of infestation varied from 0 to 21%. The propagation rate of parasitic nematodes in the haemocoel of infected fleas was very high; in some cases, we observed up to 1,000 juveniles per flea specimen. Our study of morphology, life cycle, and rDNA sequences of these parasites revealed that they belong to three distinct species differing in the host specificity. On SSU and LSU rRNA phylogenies, these species representing three genera (Rubzovinema, Psyllotylenchus, and Spilotylenchus), constitute a monophyletic group close to Allantonema and Parasitylenchus, the type genera of the families Allantonematidae and Parasitylenchidae (Nematoda: Tylenchida). We discuss the SSU-ITS1-5.8S-LSU rDNA phylogeny of the Tylenchida with a special emphasis on the suborder Hexatylina.

1. Introduction

More than 150 species of fleas feeding on different mammalian hosts, primarily rodents, are vectors of the bacterium Yersinia pestis, a causative agent of plague [1, 2]. In natural foci of plague, the dynamics of flea populations are among the main factors controlling the incidence of epizootics that pose a threat to humans inhabiting the areas [3–5]. Entomoparasitic nematodes of the order Tylenchida are known to control populations of various insect hosts [6–9]. The rate of tylenchid infestation in fleas reaches 50–60% in some cases [10, 11], when the nematodes cause castration and early death of the flea hosts [9, 12, 13].

Despite high importance of the Tylenchida as a nematode order harboring entomoparasites and notorious crop pests, their reliable phylogeny is still a challenge. Tylenchid nematodes differ widely in life cycle, parasitic strategies, and the host range that spans plants, fungi, and invertebrates. Phylogenies obtained from SSU and partial LSU rDNA data often disagree with classifications based on morphology and life cycle [14–21]. Phylogenetic resolution inside the order is far from being clear, which in many respects results from the insufficiency of data available to adequately describe its diversity. As for tylenchid parasites of fleas, only 31 species are described to date [9, 22–31], with no molecular vouchering. Here we present a study of parasitic nematodes isolated from fleas sampled from different rodent hosts in a natural focus of plague.

2. Materials and Methods

2.1. Collection of Samples

Samples were collected in 2012 (spring and autumn) and 2013 (spring) in the Volga-Ural natural focus of plague (Figure 1). The sampled rodents included sousliks (Citellus pygmaeus), mouse-like rodents (Microtus socialis and Apodemus uralensis), and jerboas (Allactaga major). Three flea species (Citellophilus tesquorum, Neopsylla setosa, and Frontopsylla semura) were sampled on sousliks; two species (Amphipsylla rossica and Ctenophthalmus secundus) were on M. socialis voles; and one species (Mesopsylla hebes) was on jerboas. Fleas were examined for nematode infestation (Table 1). Examination and dissection of fleas were carried out using the dissecting microscope MBS-2 (LOMO, Russia). A half of parasitic nematodes sampled from each flea was preserved for subsequent DNA extraction, and another half was used for morphological analysis. Live fleas infected with nematodes were placed in glass flasks with river sand to obtain free-living forms. Insects were kept in a KBF 720 (E5.2) climate chamber (Binder, Germany) at 26°C and 80% humidity.

Table 1: Number of fleas studied and the percentage of fleas infected with nematodes.

Time of sampling
Host rodent species
Flea species
Number of collected fleas
Number of infected fleas
Percentage of infected fleas

April 2012
Citellus pygmaeus
Citellophilus tesquorum
41
7
17.1%

Neopsylla setosa
73
5
6.8%

Frontopsylla semura
54
7
13%

October 2012
Microtus socialis
Amphipsylla rossica
135
9
6.7%

Ctenophthalmus secundus
88
1
1.1%

April 2013
Citellus pygmaeus
Citellophilus tesquorum
34
0
0

Neopsylla setosa
271
22
8.1%

Frontopsylla semura
19
4
21%

Microtus socialis and Apodemus uralensis
Amphipsylla rossica
6
0
0

Ctenophthalmus secundus
52
0
0

Allactaga major
Mesopsylla hebes
34
2
5.9%

Figure 1: The sampling region on the map of Europe.

2.2. Morphological Analysis

Fixation and clarification of nematode preparations were performed using standard techniques described by De Grisse [32]. Material was mounted on slides in a drop of glycerin, bound by a paraffin circlet (http://pest.cabweb.org). Color staining of preparations was not performed. Morphometric analysis was conducted using the light microscope “Leica DM 1000” (Leica, Germany) with an eyepiece micrometer. Pictures of nematodes were taken with the microscope “DFC 425” (Leica, Germany). Published data on morphometrics [23, 25, 26] were used for comparison.

2.3. DNA Extraction, PCR, and Sequencing

DNA samples were extracted with a Diatom DNA Prep (IsoGen Lab, Russia). rDNA fragments were amplified using an Encyclo PCR kit (Evrogen, Russia) and primers given in Table 2. The amplified rDNA fragments were sequenced using an Applied Biosystems 3500xL DNA analyzer. Sequence reads were assembled with the CAP contig assembly program [33] and proofread with the BioEdit software [34]. For three isolates, almost complete sequences of 18S and 28S rRNA and complete sequences of 5.8 rRNA, internal transcribed spacers ITS1 and ITS2 were assembled. The sequences were submitted to GenBank under accession nos. KF155281–KF155283. For the rest of isolates, partial (750–800 bp) sequences of 18S and 28S rRNA genes were submitted to GenBank under accession nos. KF373731–KF373740.

Table 2: Nucleotide sequences of primers used in this study.

Primer
Sequence
Orientation
References

Nik22
tmycygrttgatyctgyc
F
This study

A
gtatctggttgatcctgccagt
F
[35]

Q5nemCh
gccgcgaayggctcattayaac
F
This study

G18SU
gcttgtctcaaagattaagcc
F
[36]

Ves18-d9
gtcgtaacaaggtatccgtaggtgaac
F
This study

R18Tyl1
ggtccaagaatttcacctctc
R
[36]

B
gtaggtgaacctgcagaaggatca
R
[35]

Q39nem
gaaaccttgttacgacttttrcbygg
R
This study

58d1
rcatcgatgaagaacgywg
F
[37]

58r nem
gcwgcgttcttcatcgacyc
R
This study

28d3
gtcttgaaacacggaccaagg
F
[37]

28d6
ggtyagtcgrtcctrag
F
[37]

D2A
acaagtaccgtgagggaaagttg
F
[38]

28r4
gctatcctgagggaaacttcgg
R
[37]

28r2nem
cggtacttgttcgctatcg
R
This study

28r7
agccaatccttwtcccgaagttac
R
[37]

28r12
ttctgacttagaggcgttcag
R
[37]

D3B
tcggaaggaaccagctacta
R
[38]

2.4. Phylogenetic Analysis

The newly obtained rDNA sequences of tylenchid parasites of fleas were aligned with a selected set of other tylenchid sequences obtained from the GenBank. The main selection criterion was to sample representatives of all clades that occur in published SSU and LSU rDNA phylogenies of the Tylenchida [16–21, 39]. Apart from the D2-D3 LSU rDNA expansion segment commonly used in previous studies, we included all LSU rDNA sequence data available for the Tylenchida, with the exception of Basiria sp. SAN-2005 (accession nos. DQ145619, DQ145667) that in our preliminary analyses (data not shown) demonstrated a disputable affinity to the Tylenchida. For the species Anguina tritici, Globodera pallida, Heterodera glycines, Pratylenchus vulnus, and Radopholus similes the nearly complete rDNA sequences were assembled with appropriate cDNA fragments identified with BLAST [40]. Partial LSU rDNA sequence of Ditylenchus dipsaci was combined with the soil environmental clone NTS_28S_061A_2_b4 (accession no. KC558346), as the clone sequence appeared to represent a close tylenchid relative of D. dipsaci. Chimeric sequences were also created in some cases when closely related partial rDNA sequences were found in the database. All sequences and their accession numbers are listed in Table 3. Cephalobidae and Chambersiellidae were chosen as the outgroup. Alignments were constructed with the MUSCLE program [41] and refined manually using the MEGA 5.0 software package [42]. Three alignments were generated: (1) SSU rDNA, (2) D3 region of LSU rDNA, and (3) concatenated rDNA data including SSU, LSU, 5.8S rDNA, and highly conserved regions of ITS1. After discarding ambiguously aligned positions, the alignments length was 1,723, 592, and 4,930 positions, respectively. Bayesian reconstruction of phylogeny was done with the PhyloBayes software, version 3.2 [43] under the GTR + CAT + DP model [44]. Eight independent runs were performed with 4,000,000 cycles each; the first 3,000,000 cycles were discarded. A consensus tree with Bayesian posterior probabilities was constructed for the remained tree sample. Bayesian reconstruction was also performed using the MrBayes software [45] under the GTR + G8 + I model [46] in two independent runs, each with four Markov chains. The chains were run for 5,000,000 generations, with trees sampling every 1,000th generation. The consensus posterior probabilities were calculated after discarding the first 3,000,000 generations. Partitioning “by genes” was used for the concatenated alignment with all parameters unlinked, except for the topology and branch lengths. In addition, node support was estimated with maximum likelihood bootstrap as implemented in the RAxML software, version 7.2.6 [47], under the GTR + G + I model with 1,000 bootstrap replicates. Alternative topologies were tested using the approximately unbiased (AU) [48] and Kishino and Hasegawa [49] tests implemented in the CONSEL software [50] and the expected likelihood weight test [51] implemented in the TREE-PUZZLE software [52]. TREEVIEW [53] was used as the tree viewer and editor, and site-wise log-likelihoods were computed with TREE-PUZZLE under the GTR + G8 + I model with substitution matrix parameters estimated by MrBayes.

Table 3: List of OTUs and accession numbers of sequences.

Name
18S rRNA
ITS1-5.8S 

rRNA
28S rRNA
%, SSU-ITS1- 

5.8S-LSU/D3
Reference
Family by [8]

Chambersiellidae*

Fescia grossa
KC242218
—
DQ145636 DQ145684
87.1/—
[54]

[55]
Chambersiellidae

Geraldius sp. SAN-2010a
—
—
GU062821
17.8/—
[56]
Chambersiellidae

Cephalobidae

Acrobeloides maximus
EU196016
JX026706
EU195987
94.8/—
[57]

[58]

[57]
Cephalobidae

Cephalobus cubaensis
AF202161
AF202161
EU253570
89.8/—
[59]

[57]

Panagrolobus sp. SN-2010
—
—
HM439771
51.9/—
[60]

Cephalobidae Gen. sp.

MHMH-2008
FJ040406
—
—
Holterman et al., 2008, unpublished.

Zeldia punctata
—
DQ146426
EU195988
96.6/—
[61]

[57]

Zeldia sp.
AY284675
—
—

Aphelenchidae

Aphelenchus avenae
JQ348399
AF119048
—
96.9/—
[62]

[63]
Aphelenchidae

Aphelenchus sp.
—
—
DQ145664 DQ145714
[55]

Paraphelenchus acontioides
—
—
HQ218322
45.5/—
[64]

Paraphelenchus sp.
AY284642
—
—
[18]

Hexatylina + “Anguinata (part)”: Iotonchioidae

Allantonema mirable
—
—
JX291132
10.6/85.8
[39]
Allantonematidae

Bradynema listronoti
DQ915805

DQ915804
45.6/96.8
[65]

Bradynema rigidum

DQ328730
10.4/86.3
[20]

Contortylenchus sp.
—
—
DQ328731
—/85.4
[20]

Deladenus durus
JQ957898
—
—
34.0/—
[66]
Neotylenchidae

Deladenus proximus
JF304744
JF304744
—
35.2/—
[67]

Deladenus siricidicola isolate 354
AY633447

AY633444
45.8/98.1
[68]

Deladenus siricidicola isolate 466
FJ004890
FJ004890
—
41.7/—
[69]

Deladenus siricidicola isolate 1093
FJ004889
FJ004889
—
42.0/—
[69]

Fergusobia camaldulensae
AY589294
—
AY589346
45.7/98.0
[68]

Fergusobia sp. 444
EF011667
—
EF011675
45.7/97.3
[68]

Fergusobia sp. SBG
FJ393270
—
FJ386996
45.7/98.3
[70]

cf. Gymnotylenchus sp. TSH-2005
AY912040
—
—
12.9/—
Powers et al., unpublished.

Howardula aoronymphium
AY589304
AY589304
AY589395
49.7/96.1
[68]
Allantonematidae

Howardula dominicki
AF519234
AF519234
—
37.4/—
[71]

Howardula neocosmis
AF519226
AF519226
—
38.2/—
[71]

Howardula phyllotretae
JX291137
—
DQ328728
41.9/86.1
[39]

[20]

Howardula sp. CD353
—
—
JX291131
—/93.9
[39]

Howardula sp. SP-A
AF519232
AF519232
—
37.7/—
[71]

Howardula sp. SP-F
AF519222
AF519222
—
38.2/—
[71]

Howardula sp. SP-MA
AF519233
AF519233
—
38.1/—
[71]

Howardula sp. SP-PS
AF519231
AF519231
—
38.1/—
[71]

Parasitylenchus  bifurcatus
KC875397
—

44.0/85.3
[72]

Parasitylenchus sp.
—
—
DQ328729
[20]

Psyllotylenchus sp. ex Frontopsylla semura
KF373734
—
KF373739
27.1/93.7
This study
Parasitylenchidae

Psyllotylenchus sp. ex Neopsylla setosa
KF373733
—
KF373738
27.1/93.7
This study

Rubzovinema sp. ex Amphipsylla rossica
KF155281
KF155281
KF155281
90.0/100.0
This study
Neotylenchidae

Rubzovinema sp. ex Ctenophthalmus cecundus
KF155282
KF155282
KF155282
89.8/100.0
This study

Rubzovinema sp. ex Citellophilus tesquorum
KF155283
KF155283
KF155283
93.2/100.0
This study

Rubzovinema sp. ex Frontopsylla semura
KF373732
—
KF373737
27.1/93.7
This study

Rubzovinema sp. ex Neopsylla setosa
KF373731
—
KF373736
27.1/93.7
This study

Skarbilovinema laumondi
—
—
JX291136
10.9/91.0
[39]
Iotonchioidea

Skarbilovinema lyoni
JX291138
—
DQ328733
41.8/86.3
[39]

[20]

Spilotylenchus sp. ex Mesopsylla hebes
KF373735
—
KF373740
27.1/93.4
This study
Parasitylenchidae

cf. Sychnotylenchus sp. CSP1-09
DQ080531
—
—
12.9/—
Powers et al., unpublished.
Sychnotylenchidae

Wachekitylenchus bovieni
—
—
DQ328732
—/85.9
[20]
Parasitylenchidae

Unidentified Allantonematidae HaMW
JQ941710
—
—
18.5/—
Rhule, unpublished.
Allantonematidae

Unidentified Allantonematidae NK2011_2
AB663183
—
—
12.0/—
[73]

Unidentified Allantonematidae NK2011_3
AB663184
—
—
12.0/—
[73]

Unidentified nematode 804U-025
EU880149
—
—
12.0/—
[74]

Unidentified nematode CD289
—
—
JX291133
—/84.1
[39]

Unidentified nematode RGD591T12
AB455970
—
—
12.0/—
[73]

Unidentified nematode WY2009_BAR-1
—
—
FJ661075
—/96.3
[75]

Unidentified parasite ex Chrysobothris affinis
—
—
DQ202658
—/51.0
Hunt et al., unpublished.

Hexatylina + “Anguinata (part)”: Sphaerularioidea

Deladenus sp. PDL-2005
AJ966481
—
—
35.0/—
[16]
Neotylenchidae

cf. Helionema sp. MHMH-2008
EU669913
—
—
34.0/—
[19]
Parasitylenchidae (genera dubia in Hexatylina)

cf. Hexatylus sp. Westplace
AY912050
—
—
12.9/—
Powers et al., unpublished.
Neotylenchidae

Nothotylenchus acris
AY593914
—
—
34.0/—
[76]
Anguinidae

Sphaerularia bombi
AB250212
—
DQ328726
56.7/100.0
Takahashi, unpublished.

[20]
Sphaerulariidae

Sphaerularia vespae
AB300595
AB300595
AB300596
54.7/100.0
[77]

Unidentified nematode 801L-022
EU880129
—
—
12.1/—
[74]

Anguinata

Anguina tritici
AY593913
JF826515
HO058555 DQ328723
57.6/92.9
Holterman et al., unpublished.

Rao and Rao, unpublished.

Rao et al., unpublished.

[20]
Anguinidae

Ditylenchus adasi
EU669909
—
—
34.6/—
[19]

Ditylenchus angustus
AJ966483
—
—
34.6/—
[16]

Ditylenchus destructor

JX162205

50.0/99.5
[78]

Ditylenchus dipsaci
AY593911
AY593911
JF327759
60.9/100.0
[76]

Zhao 2011, unpublished.

clone NTS_28S_061A_2_b4

KC558346
[79]

Ditylenchus drepanocercus
JQ429768
JQ429774
JQ429772
48.7/89.3
[80]

Ditylenchus halictus
AY589297

52.8/97.3
[68]

Ficotylus congestae
EU018049

45.6/97.5
[81]

Halenchus fucicola
EU669912
—
—
34.6/—
[19]

Pseudhalenchus minutus
AY284638

34.6/—
[19]

Unidentified entomoparasitic nematode SAS-2006 “Neotylenchus ” sp.
—
—
DQ328725
—/85.6
[20]

“Tylenchina”: Tylenchidae

Aglenchus agricola
FJ969113
—
—
46.0/—
van Megen et al., unpublished.
Tylenchidae

Aglenchus sp.
—
—
JQ004996
[82]

Coslenchus costatus
AY284581
—
—
45.5/— 
[18]

Coslenchus sp.
—
—
JQ005007
[82]

Filenchus annulatus
JQ814880
—
JQ005017
46.4/—
[82]

Tylenchus davainei
AY284588
—
—
33.9/—
[18]

“Tylenchina”: Tylodoridae

Eutylenchus excretorius
EU915487
EU915500
EU915490
35.8/—
[83]
Atylenchidae

Cephalenchus hexalineatus
AY284594
—
—
44.1/—
[18]
Tylodoridae

“Tylenchina”: Boleodoridae

Basiria gracilis
EU130839
—
DQ328717
44.6/—
[84]

[20]
Tylenchidae

Basiria sp. 3 TJP-2012
—
—
JQ004998
12.0/—
[82]

Boleodorus thylactus
AY993976
—
—
46.7/— 
[16]

Boleodorus sp.
—
—
JQ005001
[18]

Neopsilenchus magnidens
AY284585
—
—
45.6/— 
[18]

Neopsilenchus sp. 3 TJP-2012
—
—
JQ005020
[82]

Neopsilenchus sp. 1 TJP-2012
—
—
JQ005018
11.9/—
[82]

“Hoplolaimina”: Merliniidae

Nagelus leptus
—
—
DQ328715
45.2/— 
[20]
Telotylenchidae

Nagelus obscurus
EU306350
—
—
[17]

Pratylenchoides ritteri
AJ966497
—
JX261964
48.7/— 
[16]

[85]
Pratylenchidae

Psilenchus cf. hilarulus
AY284593
—
EU915489
44.1/—
[18]

[83]
Psilenchidae

Scutylenchus quadrifer
AY284599
—
—
41.5/— 
[18]
Telotylenchidae

Scutylenchus sp.
—
JQ069956
—
[86]

“Tylenchina”: Ecphyadophoridae

Ecphyadophora sp. JH-2004
AY593917
—
—
33.7/—
[76]
Ecphyadophoridae

“Ditylenchus” brevicauda
AY284635
—
—
33.9/—
[18]
Anguinidae

Malenchus andrassyi
AY284587
—
—
32.3/—
[18]
Tylenchidae

Ottolenchus discrepans
AY284590
—
—
33.7/—
[18]

Criconematina

Hemicriconemoides gaddi
—
KC520471
KC520470
55.6/— 
[87]
Criconematidae

Hemicriconemoides pseudobrachyurus
AY284622
—
—
[18]

Hemicycliophora lutosa
—
GQ406237
GQ406240
53.2/— 
[88]
Hemicycliophoridae

Hemicycliophora thienemanni
AY284628
—
—
[18]

Meloidoderita kirjanovae
—
DQ768427
DQ768428
50.8/—
[89]
Sphaeronematidae

Sphaeronema alni
FJ969127
—
—
van Megen, unpublished.

Meloidoderita sp.
GU253916
GU253917
JQ771954
50.8/—
[90]

Cudejkova and Cermak, unpublished.

Tylenchulus semipenetrans
AJ966511
FJ588909
FJ969710
57.5/—
[16] 

[91] 

[92]
Tylenchulidae

“Hoplolaimina”: Belonolaimidae

Belonolaimus longicaudatus
AY633449
DQ672366
GQ896548
55.8/—
[68]

[93]

[94]
Belonolaimidae

Ibipora lolii
JQ771535
—
—
30.9/—
[95]

“Hoplolaimina”: Hoplolaimidae

Carphodorus sp.
JQ771538
—
JQ771550
41.3/—
[95]

Globodera pallida
EU855119
EU85511
BM415342

BM415248

CV577211

CV577977

CV579301E

U85511
93.6/—
Nowaczyk et al., unpublished.

Opperman, unpublished 

[96].
Heteroderidae

Heterodera glycines
AF216579

BI704127

BI748392

CA940548

CB379240

CB379263

CB379850

CB380242

CB825296

CB825409

CB825970

CB935610

CK348871

CK348904

CK349175

CK352112
AF216579
AF133304

AF216579

BI704144

BI704144

BI749520

CA940190

CA940212

CA940243

CA940406

CA940424

CA940429

CA940589

CB238697

CB279977

CB299455

CB373844

CB373981

CB379125

CB379140

CB379219

CB379312

CB379439

CB379505 

CB379696

CB379707

CB379996

CB380091

CB380241

CB824788

CB824878

CB825995

CB934877

CB934931

CB934950

CB934954

CK348525

CO036619

HM560850

JN684906
98.3/—
[97] 

[96].

[98] 

Yan and Davis, unpublished.

[99]

Ye et al., unpublished.

Wei et al.,

unpublished.

Morulaimus sp.
JQ771540
—
—
31.5/—
[95]
Belonolaimidae

Radopholus similis
AJ966502

AY912509

EF384224

EY190988

EY191076

EY191697

EY191883

EY192786

EY192788

EY193123

EY193253

EY194340

EY194464

EY194646

EY195472

FJ040398
AY912509 

EF384224
EU555409

EY189839

EY190550

EY190620

EY190961

EY191066

EY191073

EY191135

EY191160

EY191173

EY191237

EY192021

EY192028

EY192080

EY192091

EY192247

EY192381

EY192472

EY192501

EY192526

EY192892

EY192907

EY193005

EY193037

EY193249

EY193314

EY193798

EY193897

EY193971

EY194395

EY194454

EY194530

EY195146

EY195204
97.5/—
[16]

[100]

Long et al., unpublished.

[101] 

Holterman et al., unpublished.

[102] 

[100]

Zhao unpublished.

[86]
Pratylenchidae

EY195406

EY195408

EY195580

EY195889

EY195943

GQ281471

JN091962

JQ782249

Rotylenchulus reniformis
JX406356
FJ374686
HM131884

FJ906072
59.4/—
[103]

Rahman et al., unpublished.

[104]
Rotylenchulidae

“Hoplolaimina”: Pratylenchidae

Dolichodorus sp. WY-2006
DQ912918
—
—
33.9/—
[105]
Dolichodoridae

Hirschmanniella loofi
EU306353
EU620472
EU620469
51.6/—
[17]

[106]
Pratylenchidae

Macrotrophurus arbusticola
AY284595
—

33.9/—
[18]
Telotylenchidae

Meloidogyne arenaria
U42342
U42342
U42342

AF023855

AF023856
99.2/—
Georgi and Abbott, unpublished.
Meloidogynidae

Meloidogyne artiellia
AF248477
AF248477
AF248477
99.2/—
[107]

Nacobbus aberrans
AJ966494
DQ017473
U47557
49.0/—
[16]

[108]

[109]
Pratylenchidae

Pratylenchus vulnus
EU669955
JQ966892
BQ580554

CV198923

CV198995

CV199233

CV199349

CV199490

CV200136

CV200423

CV200464

CV200467

CV200471

CV200530

CV200687

CV200896

CV201004

CV201135

EL887566

EL887705

EL888035

EL888060

EL888174

EL888269

EL888739

EL888778

EL889241

EL889472

EL889797

EL889934

EL889934

EL889977

EL889994

EL890380

EL890701

JQ003993

JQ003994

JX047008
100.0/—
[19]

[110] 

[96] 

[96]

[111]

Zhao, unpublished.

[112]

Tylenchorhynchus dubius
EU306352
—
DQ328707
53.2/—
[17]

[20]
Telotylenchidae

Tylenchorhynchus zeae
—
EF519711
—
[113]

Clades of the tree, marked by boldface.

3. Results

3.1. Infestation of Fleas with Nematodes

The infestation rate is shown in Table 1 (in total, 807 flea specimens were studied). Among the six flea species studied, the population size and the percentage of infected fleas varied depending on the season. Three flea species sampled on sousliks (Citellophilus tesquorum, Neopsylla setosa, and Frontopsylla semura) exhibited a stable population density. In the two species, N. setosa and F. semura, the infestation rate was moderate to high in the spring seasons of 2012 and 2013. In C. tesquorum, no infected fleas were detected in spring 2013, whereas in spring 2012 the fleas were highly infested (17.1%). The vole flea Amphipsylla rossica was abundant and moderately infested in autumn, whereas being less abundant in spring, which may explain the absence of infected fleas in the spring sample. Another vole flea, Ctenophthalmus secundus, exhibited a consistently high population density and low infestation rate in both spring and autumn samples.

Adult parasitic females and their progeny were found in the haemocoel of infected fleas. In the infected fleas C. tesquorum, A. rossica, C. secundus, and Mesopsylla hebes, only one generation of parasitic females was observed. Their amount in a flea specimen is determined by the number of free-living infective females that penetrate into the flea larva. We observed 1 to 2 or 1 to 4 adult parasitic females per flea specimen in spring and autumn, respectively. An additional parthenogenetic generation of parasitic females was found in some fleas of N. setosa and F. semura, where up to 16 specimens per flea were observed. As in other entomoparasitic nematodes, the propagation rate depends on the host age. Thus, in young fleas up to 10 juveniles was found per flea specimen, whereas up to 1,000 juveniles of different stages were contained in some old fleas (Figure 2). After the 2nd molt the number of juveniles is maximal, and 3rd stage juveniles massively migrate to the rectal section of the flea intestine for exit to the environment. In some cases, the observed infestation level was so high that nematodes penetrated distal segments of the flea legs, from where they have no way to the environment.

Figure 2: Numerous juveniles of Rubzovinema sp. extracted from the dissected body of a Citellophilus tesquorum flea.

3.2. Morphological Analysis of Entomoparasitic Stages in Nematode Isolates and Their Taxonomic Identification

Analysis of morphology of entomoparasitic stages suggests that the studied nematode isolates from three distinct groups. A single generation of parasitic females was observed in the first two groups and an additional parthenogenetic generation—in the third group. According to morphometric data on adult parasitic females (Tables 4–6), the first two groups belong to the genera Rubzovinema or Spilotylenchus and the third group to the genus Psyllotylenchus. Photographs of parasitic females of Rubzovinema sp., Spilotylenchus sp., and Psyllotylenchus sp. are depicted in Figure 3. Figure 4 shows their distribution among flea samples studied.

Table 4: Comparison of morphometrics in parasitic females of Rubzovinema sp. and Rubzovinema  ceratophylla.

Character
Rubzovinema sp. (this study)
Rubzovinema ceratophylla [26]

N
29
27

L
1278,6 (840–1570)
1265,1 (810–1840)

D
120,8 (85–145)
137,3 (62–200)

A
11,19 (7,9–16,1)
9,51 (6,4–16,8)

C
65,4 (31,4–100)
44,10 (10–86,4)

V%
96,4 (93,1–97,9)
95,44 (92–98,9)

Total length of stylet (St)
18,5 (14–22)
19,5 (18–21)

Length of distal edge of stylet
7,2 (5–8,7)
—

Distance between anterior end and excretory pore (Ex)
20,7 (10–31)
—

Distance between anterior end and nerve ring
61,2 (50–74,5)

Total length of tail (Cd)
21,9 (10–42)
26,35 (14–47,5)

Distance between vulva and tail end
46,1 (23–75)
—

Distance between vulva and anus (V–A)
26,9 (13–40)
—

All measurements are in m and in the form mean (range).

Table 5: Comparison of morphometrics of parasitic females in Spilotylenchus sp. and Spilotylenchus maisonabei.

Characters
Spilotylenchus sp. (this study)
Spilotylenchus maisonabei [23]

N
2
6

L
1,600–1,840
1,244 (1,200–1,320)

D
155–160
125 (107–160)

A
10.3–11.5
10.3 (7.5–12)

C
167.3–177.8
84.4 (64.5–121)

V%
97.4–97.7
96.2 (95.8–96.5)

Total length of stylet (St)
9.5–9.8
9-10

Distance between anterior end and excretory pore
1.5–15.5
23.3 (20–28)

Distance between anterior end and nerve ring
—
52–54

Total length of tail (Cd)
9–11
15.4 (10–19)

Distance between vulva and tail end
41.5–43
47 (42–52)

Distance between vulva and anus (V–A)
32-33
—

All measurements are in m and in the form mean (range).

Table 6: Comparison of morphometrics of parasitic females in Psyllotylenchus sp. and Psyllotylenchus viviparous.

Character
Psyllotylenchus sp. (this study)
Psyllotylenchus viviparous [25]

Gamogenetic
Parthenogenetic
Gamogenetic
Parthenogenetic

N
3
7
8
10

L
1,016.7 (900–1,100)
446 (420–500)
1,000 (840–1,480)
500 (360–840)

D
81.3 (79–84)
70 (60–80)
77 (62–115)
60 (54–100)

A
12.5 (11.1–13.3)
6.25 (5.6–7)
—
—

C
64.3 (60–68.2)
40.15 (37.1–43.5)
—
—

V%
95.1 (95–95.4)
93.3 (90–95.3)
—
—

Total length of stylet (St)
17.5 (17–18,5)
5.25 (4–6)
17 (15–20)
7 (5–8)

Length of the distal edge of stylet
8.6 (8-9)
—
—
—

Distance between anterior end and excretory pore
26.5 (25–31.5)
17.5 (15–19.5)
23 (13–33)
22 (14–46)

Distance between anterior end and nerve ring
—
51.7 (50–55)
—
—

Total length of tail (Cd)
15.8 (15–17)
11.1 (10.5–11.5)
25 (17–35)
9 (1–17)

Distance between vulva and tail end
48 (45–51)
30.5 (19.7–55)
56 (37–71)
52 (40–104)

Distance between vulva and anus (V–A)
30.8 (29–31.5)
13.5 (11.7–21.6)
—
—

All measurements are in m and in the form mean (range).

(a)

(b)

(c)

(d)

Figure 3: Parasitic females of the studied nematode species. (a) Rubzovinema sp., heterogeneous female; (b) Spilotylenchus sp., heterogeneous female; (c) Psillotylenchus sp., heterogeneous female of the first generation; (d) (c): Psillotylenchus sp., parthenogenetic female of the second generation. Scale bar—200 μm.

Figure 4: Distribution of the studied nematode species among the flea species studied, whose rodent hosts are given below. The vertical axis shows the numbers of infected fleas.

According to morphometric evidence, parasitic females and juveniles of the genera Rubzovinema and Spilotylenchus are very similar. However, in the first two groups of isolates we found characters bearing discriminative and identificational value. In particular, the oesophageal glands in juveniles III of the first group are poorly developed. This is a distinctive feature of the genus Rubzovinema, where males and females have shortened oesophageal glands located close to the nerve ring. In the second group of isolates, oesophageal glands are well developed and elongated, which is characteristic of the genus Spilotylenchus. In the first group, the stylet possesses a heavily sclerotized distal spear with a length of approximately half the total stylet length and has a stem with a weaker sclerotization and widening to the base. This stylet structure is characteristic of the genus Rubzovinema, and stylet length (18.5 (14–22) μm) is in accordance with morphometrics given in the description of this genus [26]. In the genus Spilotylenchus, the stylet varies in shape but always possesses a shortened conical distal spear. In the second group of isolates, the stylet structure was similar to that of Spilotylenchus. Also, the vulval lips of the first group are more protruded than in Spilotylenchus. Other features, including the morphometrics, vary widely in both genera, which hampers taxonomic identification. Nevertheless, based on distinctive traits, we identified the first and second group of isolates as Rubzovinema sp. and Spilotylenchus sp., respectively.

In the genus Rubzovinema, the single species described to date is Rubzovinema ceratophylla [26]. This species is known to parasitize exclusively the flea Citellophilus tesquorum that feeds on sousliks. The specimens of Rubzovinema studied in this work were isolated from five flea species, C. tesquorum, Neopsylla setosa, Frontopsylla semura, Amphipsylla rossica, and Ctenophthalmus secundus, of which the latter two were sampled on mouse-like rodents. Also, the parasitic females of Rubzovinema sp. differed from R. ceratophylla by morphology; they have a shorter tail and more protruded vulval lips. A morphometric comparison of Rubzovinema sp. and R. ceratophylla is given in Table 4.

The parasitic females of Spilotylenchus sp. were isolated from the flea Mesopsylla hebes associated with jerboas. The females were not identified to the species level because of a small number of available specimens and the lack of a free-living stage. A morphometric comparison of Spilotylenchus sp. and the morphologically closest species Spilotylenchus maisonabei [23] is given in Table 5.

In the genus Psyllotylenchus, descriptions of most species are fragmentary and incomplete, which precluded the species identification of the Psyllotylenchus isolates from the fleas N. setosa and F. semura feeding on sousliks. A morphometric comparison of Psyllotylenchus sp. and the type species of this genus, Psyllotylenchus viviparous [25], is given in Table 6.

The 18S and 28S rDNA sequences of Rubzovinema sp. specimens from A. rossica and C. secundus were 100% identical, which indicates that the isolates belong to the same species. The sequences of Rubzovinema sp. ex C. tesquorum, Rubzovinema sp. ex N. setosa, and Rubzovinema sp. ex F. semura diverged from one another and from the gene sequences of Rubzovinema sp. ex A. rossica and Rubzovinema sp. ex C. secundus by 0.4–0.7%, which corresponds to the levels of intraspecific variation [14, 114–119]. The 18S and 28S rDNA sequences of Psyllotylenchus sp. ex N. setosa and Psyllotylenchus sp. ex F. semura were 100% identical, indicating that they belong to the same species. The 18S and 28S rDNA sequences of Rubzovinema sp. and Psyllotylenchus sp. diverge by 1.2% and 1.9%, respectively. Those of Spilotylenchus sp. ex M. hebes were found to be more divergent. The degree of divergence of the 18S rDNA sequence of Spilotylenchus sp. ex M. hebes from those of either Rubzovinema sp. or Psyllotylenchus sp. was 2.4%; the D3 expansion segment of 28S rDNA diverged by 13.1% and 12.0%, respectively. The observed divergence rate of rDNA sequences agrees well with published evidence on entomoparasitic nematodes [14, 114–118]. Thus, intraspecific divergence of 18S rDNA in Deladenus siricidicola is 1% [120], of D2 and D3 expansion segments in the phytoparasite Bursaphelenchus xylophilus is from 0% to 0.6%, and the interspecific variation between the phytoparasites B. xylophilus and Bursaphelenchus mucronatus is from 1.7% to 3.7%. The spacers ITS1 and ITS2 are generally more diverged; the intra- and interspecific variation for these species is from 0 to 3.1% and 11.2 to 13.4%, respectively [121–123].

Molecular vouchering is proved to efficiently complement morphological species identification in nematodes [73, 122, 124–128]. Combining the rDNA and morphological data confirms the species identity within each of the three studied groups of isolates.

3.3. Phylogenetic Analysis

In phylogenetic analyses of rDNA we used a dataset with extensive species and gene sampling (SSU-ITS1-5.8S-LSU) compared to earlier published tylenchid phylogenies, most of which were based on SSU rDNA or D2-D3 expansion segments [17, 19–21, 39, 129]. The SSU-ITS1-5.8S-LSU rDNA tree topology (Figure 5) is highly similar to other published phylogenies of tylenchids. In this tree, tylenchomorphs are represented by the sister groups Aphelenchidae and Tylenchida. Most of the tylenchid clades occur in published trees but often contradict classifications based on morphology, as it was also noted by other authors [17, 19–21, 39, 129]. The three robust major branches in the SSU-ITS1-5.8S-LSU rDNA tree (Bayesian posterior probabilities of 0.99–1.0) are (1) the clade includes representatives of the suborders Hoplolaimina, Criconematina, and Tylenchina (excluding Anguinoidea); (2) the majority of classic Anguinata; (3) the suborder Hexatylina. The studied parasites of fleas form a monophyletic group (bootstrap support of 100%) within the Hexatylina.

Figure 5: Phylogenetic tree of Tylenchida, inferred from SSU-ITS1-5.8S-LSU rDNA sequences. Topology was inferred using the PhyloBayes software (maxdiff = 0.36). Node support values are shown as follows: the first two values are Bayesian posterior probability assessed using the PhyloBayes and MrBayes software, respectively, and the third is bootstrap support assessed by the ML method. Thick lines lead to the nodes, in which at least one support value of posterior probability is 0.95 and higher. Names of clades (framed) are mainly given by type genera included in them (with the exception of Iotonchioidea). Formal taxonomic position (family by [8]) is shown on the right to the color bar. Colors indicate the ecologies (see the legend). Names of the species of Hexatylina that have a mycetophagous stage in their life cycle are shown in blue. The three robust major branches of Tylenchida are marked by gradient.

The nonredundant rDNA data on the Hexatylina in GenBank mostly represents the D2-D3 expansion segments of LSU rDNA. To maximize species sampling of the Hexatylina, we chose the D3 expansion segment as the molecular marker. The phylogenetic tree with the Anguinoidea as an outgroup is shown in Figure 6. In this tree, the suborder Hexatylina consists of two well-supported clades, in accordance with previously published D2-D3 rDNA phylogenies [19, 20, 39]. The clade of the studied flea parasites is placed within the largest branch of the Hexatylina, similarly to the result of the concatenated rDNA analysis.

Figure 6: Phylogenetic tree of Hexatylina, inferred from D3 expansion segment of LSU rDNA. Topology was inferred using the PhyloBayes software. Node support values are shown as follows: Bayesian posterior probability/bootstrap support assessed by the ML method. Thick lines indicate the nodes supported at the level of 0.95 and higher. Color of lines indicates the ecologies (see the legend). Names of species were shown in different colors indicating their taxonomic position. Three families that include their type genera (shown as circles) are marked by gradient.

The three alternative relationships between the three major branches of Tylenchida (Figure 5) are not discriminated by the AU and Kishino and Hasegawa tests, and only the basal position of the Hexatylina is rejected by the expected-likelihood weights test (Table 7). All three tests do not discriminate between the alternative placement of the flea parasites as closest to the Allantonema, Parasitylenchus, or Deladenus branches; however, its positioning outside this grouping is not rejected only by a less conservative Shimodaira-Hasegawa test [50].

Table 7: Results of tree topology tests for alternative hypotheses on () the initial divergence of Tylenchida (Figure 4) and on () the relationships within the monophyletic branch that includes the studied group of nematodes parasitizing fleas (designated by asterisk).

Topology
Rank
obs
au
np
bp
pp
kh
sh
c-ELW

1

(((H,An),T),o)
1
−1.8
0.787
0.415
0.402
0.804
0.663
0.969
0.4197

((An,(H,T)),o)
2
4.1
0.326
0.198
0.205
0.013
0.254
0.623
0.1848

((H,(An,T)),o)
3
6.9
0.061
0.013
0.014
0.001
0.101
0.492
0.0186

2

((((*,Al),P),Ds),o)
1
−1.8
0.787
0.415
0.402
0.804
0.663
0.969
0.4197

((((*,P),Al),Ds),o)
2
1.8
0.495
0.242
0.247
0.130
0.337
0.813
0.2249

(((*,(Al,P)),Ds),o)
3
2.7
0.371
0.110
0.105
0.052
0.243
0.824
0.1209

((*,((Al,P),Ds)),o)
6
15.7
0.063
0.024
0.025

0.053
0.153
0.0272

(((*,Ds),(Al,P)),o)
7
18.3
0.013
0.002
0.002

0.020
0.096
0.0028

Al: Allantonematidae, An: Anguinata, Ds: Deladenus siricidicola —D. proximus group, H: Hexatylina, P: Parasitylenchidae, T: Tylenchina, o: outgroup.

4. Discussion

4.1. Ribosomal DNA Phylogeny of the Tylenchida and Relationships within the Suborder Hexatylina

Phylogenetic analyses of SSU [16, 17, 19, 39] and D2-D3 [20, 39] rDNA data using various methods and species sampling generally agree on the monophyly of most tylenchid clades and contradict classic morphology based classifications. In the SSU-ITS1-5.8S-LSU tree (Figure 5), the monophyletic Tylenchida consists of three major robust clades. The first clade diverges into six groups: (1) the “Tylenchidae (part 2)” (by [17]), (2) the Tylodoridae (represented by the two genera, Cephalenchus and Eutylenchus [83]), (3) Boleodorinae + “Tylenchidae (part 1)” (by [Bert]), (4) the Merliniidae [130], (5) Criconematina + Sphaeronematidae + selected Tylenchina, and (6) Belonolaimidae + “Hoplolaimina.” The Merliniidae group corresponds to Clade C in [19] and includes partially the polyphyletic “Telotylenchinae” [131], “Pratylenchidae”, and “Hoplolaimina” (Psilenchus cf. hilarulus). Group (5) corresponds to Clade 12A in [129], where Sphaeronematidae (Sphaeronema and Meloidoderita) were earlier shown to be closely related to Criconematina [20, 89], and selected Ecphyadophoridae + Ottolenchus + Malenchus were found to represent a monophyletic clade within the paraphyletic Tylenchina likely to be related to the Criconematina [18, 82]. Group (6) corresponds to Clade VII in [20], Clade 12B in [129], and Clade A + Clade B in [19]. Belonolaimidae (the genera Belonolaimus and Ibipora) tend to occupy the basal position. Clade A in [19] contains a “long branch” of the burrowing nematode Radopholus similes (“Pratylenchidae”) in sister position to the Hoplolaimidae [17, 19]. This nematode occupies a similar position relative to the Hoplolaimidae in the SSU-ITS1-5.8S-LSU tree, and we consider this unlikely to be an LBA artefact. Similarly to [95], Carphodorus and Morulaimus that belong to the classic Belonolaimidae comprise the basal branch of Clade A sensu [19]. The clade corresponding to Clade B in [19] contains Meloidogynidae, Dolichodoridae, paraphyletic Pratylenchidae, and a part of Telotylenchidae.

The second major clade of the Tylenchida includes representatives of the classic infraorder Anguinata, with a well-supported monophyletic origin, except for a few species. They belong outside the second clade and may initially have been wrongly identified.

The third major clade includes representatives of the classic suborder Hexatylina and consists of two groups. The smaller one unites the three species of Sphaerularia, Helionema sp., cf. Hexatylus sp., Deladenus sp. PDL-2005, and Nothotylenchus acris (Anguinata: Nothotylenchidae). It is further referred to as the Sphaerularioidea according to the type genus. The larger group contains the clade of studied flea parasites and members of the superfamilies Iotonchioidea (Skarbilovinema spp., Parasitylenchus spp., and Wachekitylenchus bovieni) and Sphaerularioidea (Allantonema mirable, Bradynema spp., Howardula spp., and Contortylenchus sp. (fam. Allantonematidae); Deladenus durus, Deladenus proximus, Deladenus siricidicola, Fergusobia spp., and Gymnotylenchus sp. (fam. Neotylenchidae)). One species of the Anguinata, Sychnotylenchus sp., also joins the larger group. Our study renders the genera Howardula and Deladenus paraphyletic, as was earlier shown in [19, 39, 71, 119].

The genus Howardula is paraphyletic in published rDNA and mitochondrial COI phylogenies [71]. Such characters of Howardula as the degeneration of oesophagus, tail shape, and the absence of stylet in males seem to have evolved independently by convergence. The paraphyletic genus Deladenus is more closely related to either ancestral forms of the Hexatylina or forms typical to the Anguinata. The infraorder Anguinata includes soil-dwelling nematodes, mostly mycetophagous or parasitizing various parts of plants. However, an unidentified entomoparasitic nematode was also grouped within the Anguinoidea [39]. The life cycle of Deladenus spp. is an irregular alternation of free-living and entomoparasitic forms. The nematode D. siricidicola is able of producing an unlimited number of free-living generations in the absence of the host larvae of siricid pine-killing wood wasps [132]. Like in Anguinata, the free-living forms of Deladenus spp. are fungal feeding. Such characters of Deladenus asthe mycetophagy, enlargement of subventral glands in entomoparasitic females versus their reduction in free-living forms, the hypertrophy of dorsal glands, and stylet reduction in free-living forms seem to be symplesiomorphic. Resemblance with the Anguinata is also typical of other mycetophagous free-living forms: Hexatylus (Neotylenchidae), Rubzovinema (Neotylenchidae), Prothallonema (Sphaerularioidae) Helionema (Hexatylina dubia), and Paurodontidae. For the latter, the entomoparasitic stage is expected but has never been observed. The relationship between the Hexatylina and Anguinata was earlier hypothesized based on morphology [7, 8, 130, 133, 134]. On rDNA phylogenies of tylenchids, the monophyly of the Hexatylina + Anguinata is either supported [19] or not rejected [20]. In the SSU-ITS1-5.8S-LSUrDNA tree obtained in this study, the monophyly of the Hexatylina + Anguinata has the Bayesian posterior probability of 0.91, but the maximum-likelihood bootstrap support is low; the AU and Kishino and Hasegawa tests did not discriminate between alternative hypotheses.

According to our SSU-ITS1-5.8S-LSU rDNA phylogeny (Figure 5), the major robust branches of the Tylenchida are incongruent with morphology-based classifications suggesting three rather than four suborders (the rank is adopted from morphological systems of tylenchids). Among them, the Hexatylina and Anguinata (both are monophyletic) are likely to be sister groups. The third emerged suborder includes representatives of three classic suborders: Tylenchina, Hoplolaimina, and Criconematina, among which only the latter does not contradict morphology-based classifications.

Considering ecological traits coded in Figure 5, the mycetophagy and/or facultative ectophytoparasitism are likely to be ancestral in the Tylenchida. Sedentary phytoparasites (root-knot species of Meloidogyne, the false root-knot genus Nacobbus, and cyst-forming Heterodera and Globodera) and other obligate endoparasites of plants evolved several times from free-living or facultative sedentary forms, as it was previously hypothesized in accordance with the concept of evolutionary trend to endoparasitism in phytonematodes [135]. Similarly, obligate endoparasites of insects from the Hexatylina are likely to have evolved from mycetophagous forms, with some species retaining the ancestral mycetophageous stage in the life cycle (e.g., species of the paraphyletic genus Deladenus and flea nematodes of the genus Rubzovinema). An interesting specific case in the Hexatylina is the genus Fergusobia that includes plant parasites associated with insects [68, 70], which may have transited to plant parasitism via entomoparasitism [39].

4.2. Ribosomal DNA Phylogeny of the Flea Nematodes and Their Classification

The nematodes of fleas do not group with the families known as their relatives in morphology-based systems, as these families do not form monophyletic groups in the tree. However, they do group with both type genera of the families Parasitylenchidae and Allantonematidae (Parasitylenchus and Allantonema, resp.). This grouping is preceded by a successive divergence of Deladenus durus and Deladenus siricidicola (Figure 5). As mentioned above, the pronounced free-living form in Deladenus seems to be ancestral to this group.

Only 31 tylenchid species that parasitize in fleas have been described to date. They differ by morphology, life cycle, and the host specificity, and belong to the five genera: Spilotylenchus (8 species), Psyllotylenchus (20 species), Incurvinema (1 species) Kurochkinitylenchus (1 species), and Rubzovinema (1 species). According to the classification of Siddiqi [8], the genera Spilotylenchus and Psyllotylenchus belong to the family Parasitylenchidae, whereas the genus Rubzovinema is a member of the Neotylenchidae. The two families represent two superfamilies, Iotonchioidea and Sphaerularioidea, respectively. All rDNA phylogenies published to date suggest that these superfamilies are paraphyletic [19, 20, 39], which is also inferred in our study with an extensive gene and taxon sampling.

A high degree of rDNA similarity in the three studied species suggests a closer relationship of these species than that assumed by the accepted system of classification. Earlier, Slobodyanyuk proposed to unite all known flea parasites into one family, the Spilotylenchidae. Its four subfamilies, Spilotylenchinae, Rubzovinematinae, Psyllotylenchinae, and Kurochkinitylenchinae, are discriminated based on the life cycle features [28]. In Spilotylenchinae and Rubzovinematinae, the entomoparasitic stage is represented by parasitic females of one heterosexual generation. In Psyllotylenchinae, in addition to the heterosexual generation, a parthenogenetic generation occurs in the flea haemocoel. In Kurochkinitylenchinae, two heterosexual generations exist in the haemocoel: the first generation produces parasitic females and the second generation produces both females and males [28]. Siddiqi also considered the unification of all flea tylenchids into one family but observed the need for further evidence in support [8].

Our results strongly suggest the inclusion of the three genera, Rubzovinema, Psyllotylenchus, and Spilotylenchus, in one family, the Spilotylenchidae [28]. The ribosomal DNA genetic distance within the family Spilotylenchidae is much smaller than that of certain tylenchid genera, for example, Meloidogyne (Figure 4) or Pratylenchus [19, 84].

4.3. Host Specificity of Flea Nematodes

The majority of tylenchid nematodes are monoxenous or oligoxenous; in particular, flea parasites were thought to be strictly host specific. Earlier papers suggested the lack of strict host specificity in Psyllotylenchus pawlowskyi and Psyllotylenchus viviparous [13, 25]. However, later these species were found to be heterogeneous and sustained revision [9, 27–29]. Spilotylenchus pawlowskyi and Spilotylenchus caspius were referred to as single-host parasites of the flea Coptopsylla lamellifer [27, 136]. Kurochkinitylenchus laevicepsi and Spilotylenchus ivashkini also share the same flea host, Nosopsyllus laeviceps [28, 29]. Before our study, the genus Rubzovinema was known to contain a single species, Rubzovinema ceratophylla, which parasitizes exclusively the flea Citellophilus tesquorum.

We found that at least two out of the three studied species are not single-host parasites. Psyllotylenchus sp. was shown to parasitize two flea species feeding on sousliks, Frontopsylla semura and Neopsylla setosa. Rubzovinema sp. was found on five flea species feeding on different rodent hosts: C. tesquorum, F. semura, N. setosa (all sampled from sousliks), Ctenophtalamus secundus, and Amphipsylla rossica (all sampled from voles). A. rossica, F. semura, and C. tesquorum belong to different families of the superfamily Ceratophylloidea (Leptopsyllidae and Ceratophyllidae), whereas C. secundus and N. setosa belong to the superfamily Hystrichopsylloidea. Unlike the host-specific R. ceratophylla, the studied Rubzovinema sp. parasitizes taxonomically distant fleas feeding on different rodents. Thus, the common opinion that flea nematodes are strictly host specific should be revisited.

As the two species of Rubzovinema demonstrate, even closely related parasites may exhibit different host range size. Among other known examples are the entomoparasitic nematodes of the genus Howardula parasitizing various beetles and flies [71, 137, 138], many phytonematodes [8], sibling species of parasitoid flies [128], and herbivorous insects [139]. The host range of parasites is an indicator of their evolutionary strategy in the ecosystem. Multihost parasites can be considered ecological generalists, in contrast to specialists that coevolve with a particular host. Generalists and specialists play different roles in the ecosystem [140], where they keep in balance, taking advantages and disadvantages of the two strategies. The advantages of generalization are yet to be explained by evolutionary biologists, whereas advantages of specialization are obvious, and it is generally accepted that evolution favors specialism [141, 142]. In the flea parasites, this trend is demonstrated by a greater species diversity of ecological specialists, the genera Spilotylenchus and Psyllotylenchus.

Nevertheless, the generalist Rubzovinema sp. was most abundant in the studied samples, which indicates that extending the host range may be evolutionarily successful. Besides the need to combat the immune response of several hosts, which is a requirement to widen the hosts range [143], the free-living stage of Rubzovinema sp. is to adapt to diverse microbioclimatic conditions of complex environments of rodent habitats. Multihost parasites pay a cost of adapting to alternative conditions [141, 144] compensated by stable survival of the species. Considering the spatial and temporal dynamics of flea populations feeding on a particular rodent host (one or two flea species usually dominate over a sampling season), multihost nematode parasites gain an advantage of their relative independence of population waves of either flea hosts or their rodent hosts. A higher infestation rate observed for Rubzovinema sp., compared to the two other studied species, may be an indicator of a greater ecological plasticity of this multihost parasite.

4.4. Entomoparasitic Nematodes in Natural Foci of Plague

In natural foci of plague, the epizootic dynamics are influenced by numerous climatic and biotic factors. The spatial and temporal population dynamics of the plague agent, Y. pestis, affect the population dynamics of the flea vectors and their mammalian hosts. Members of the transmission route of the plague agent also closely interact with other living organisms. For example, parasites of fleas that in turn feed on rodents are hyperparasites that play the role of high-level control agents on the ecosystem level, the role that entomoparasitic nematodes share with the bacterial plague agent. High-level control agents render the epidemiological state of a natural focus of disease less predictable. On the one hand, a lower density of the flea vector population reduces the plaque transmission rate; on the other, its growth causes an exponential decay of the host rodent population [145] below its epidemiological threshold, above which there is a threat of spillover of plague infection into human population [145]. Hypothetically, nematode-induced decrease of flea population is able to increase the number of rodents above the threshold and thus trigger an epidemic. The dual effect of high-level control agents is well exemplified by cases, when during plague episodes the extermination of rodents by humans causes the return of infection through stimulating the migration of fleas, the plaque vectors [5].

The studied entomoparasitic nematodes possess high potential as control agents of the flea vectors of plague owing to their high propagation rate within the flea host (Figure 2) and high infestation level (up to 21% observed in this study and from 50 to 60%, as estimated by other authors [10, 11]). One of the studied nematode species, Rubzovinema sp., is a multihost parasite. Host-specific parasites reach the optimal level of pathogenicity by maintaining the trade-off between pathogenicity and transmissibility. Adding of a new host to a multihost system makes the model more complicated [141]. The multihost parasite Rubzovinema sp. is expected to exhibit different levels of pathogenicity with respect to different flea hosts which, in turn, play different roles in the transmission of plague. Epizootics cause sporadic mortality in local populations of all members involved in the interaction with the plague agent, and their survival is contingent on migrations within a metapopulation. It is the case when the Cope’s law [139, 146] governs the extinction of specialists on a shorter time scale rather than a geological period, and evolution may favor the ecological generalists, such as Rubzovinema sp.

Some authors surmised the involvement of entomoparasitic nematodes in the transmission of the plague agent [4], as it was observed that biofilms of Yersinia pestis adhere to cuticle receptors of Caenorhabditis elegans [147–149]. In this perspective, nematodes parasitizing fleas in natural foci of plague take on greater importance, as they may provide for the transmission route that does not include a mammal [4]. Further studies will clarify the role of flea nematodes in the transmission of plague infection.

Conflict of Interests

The authors declare that there is no conflict of interests regarding the publication of this paper.

Acknowledgments

The authors thank G. S. Mirzaeva for help with PCR amplification and rDNA fragment analysis, N. V. Popov and the staff of the Laboratory of Epizootics Monitoring for advice and assistance in collecting and processing rodent samples, and, particularly, A. N. Porshakov for help in identification of flea specimens. They also thank O. V. Slobodyanyuk for helpful discussions of results, S. E. Spiridonov for advice on cultivation of entomoparasitic nematodes, S. A. Subbotin for valuable comments on the earlier version of the paper, and E. Yu. Talanova and L. Yu. Rusin for discussions of its final version, and E. A. Musatkina for assistance with the manuscript preparation. They are grateful to the Supercomputer Center of Moscow State University (http://parallel.ru/cluster) and the Bioportal of the University of Oslo (http://www.bioportal.uio.no) for providing computing resources.

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